Nucleic Acids Research Advance Access published May 31, 2012
Nucleic Acids Research, 2012, 1–12
doi:10.1093/nar/gks392
Requirement of upstream Hfq-binding (ARN)x
elements in glmS and the Hfq C-terminal region for
GlmS upregulation by sRNAs GlmZ and GlmYy
Nilshad N. Salim, Martha A. Faner, Jane A. Philip and Andrew L. Feig*
Department of Chemistry, Wayne State University, Detroit, MI 48202, USA
Received January 4, 2012; Revised March 27, 2012; Accepted April 13, 2012
ABSTRACT
INTRODUCTION
Small non-coding RNAs (sRNAs) are used to modulate a
wide variety of cellular responses in both bacterial and
eukaryotic systems (1–4). In eukaryotes microRNAs and
siRNAs (21 nt) often regulate translation and RNA
turnover and defend the genome from invasive nucleic
*To whom correspondence should be addressed. Tel: +1 313 577 9229; Fax: +1 313 577 8822; Email: [email protected]
yThis article looks at the mechanism through which small non-coding RNAs regulate bacterial gene expression. These pathways control stress
response and regulate expression of virulence factors necessary for bacterial pathogenesis. In this work, we dissect how an unusual pair of sRNAs
work together to control an essential gene in Escherichia coli required to make a precursor used in the cell wall biosynthesis.
ß The Author(s) 2012. Published by Oxford University Press.
This is an Open Access article distributed under the terms of the Creative Commons Attribution Non-Commercial License (http://creativecommons.org/licenses/
by-nc/3.0), which permits unrestricted non-commercial use, distribution, and reproduction in any medium, provided the original work is properly cited.
Downloaded from http://nar.oxfordjournals.org/ at Wayne State University on June 4, 2012
Hfq is an important RNA-binding protein that helps
bacteria adapt to stress. Its primary function is to
promote pairing between trans-acting small
non-coding RNAs (sRNAs) and their target mRNAs.
Identification of essential Hfq-binding motifs in
up-stream regions of rpoS and fhlA led us to ask
the question whether these elements are a
common occurrence among other Hfq-dependent
mRNAs as well. Here, we confirm the presence
of a similar (ARN)x motif in glmS RNA, a gene
controlled by two sRNAs (GlmZ and GlmY) in an
Hfq-dependent manner. GlmZ represents a canonical sRNA:mRNA pairing system, whereas GlmY is
non-canonical, interfacing with the RNA processing
protein YhbJ. We show that glmS interacts with both
Hfq-binding surfaces in the absence of sRNAs. Even
though two (ARN)x motifs are present, using a
glmS:gfp fusion system, we determined that only
one specific (ARN)x element is essential for regulation. Furthermore, we show that residues 66–72
in the C-terminal extension of Escherichia coli Hfq
are essential for activation of GlmS expression by
GlmY, but not with GlmZ. This result shows that
the C-terminal extension of Hfq may be required
for some forms of non-canonical sRNA regulation
involving ancillary components such as additional
RNAs or proteins.
acids (3). Similarly, in bacterial systems, two main
classes of sRNAs have been identified based on the
origin of the sRNAs relative to their mRNA target. Cisacting sRNAs base pair extensively with mRNAs as they
are transcribed from the opposite strand of their target
RNA (4). These systems are often used by mobile
genetic elements such as bacteriophages, transposons
and plasmids to regulate translation, RNA processing
and RNA decay (5,6). Riboswitches represent another
ubiquitous class of cis-acting RNAs. Riboswitches
respond to cellular stimuli by refolding the mRNAs via
their 50 -UTRs to trigger translation regulation or transcription termination (7). In contrast, trans-acting RNAs
are synthesized from different genetic loci than their
targets and are widely used to modulate stress responses
in bacteria. Unlike cis-RNAs, trans-acting RNAs can accommodate imperfect base pairing with their target RNAs
and often require the RNA-binding protein Hfq to facilitate complex formation.
Hfq was first identified as a host factor involved in
the replication of phage Qb RNA (8). Hfq belongs to
the Sm/Lsm family of proteins and associates into a
homo-hexameric structure to form the functional nucleic
acid-binding core. In general, the nucleic acid-binding
Sm1 and Sm2 domains are highly conserved across bacterial species (7–66 amino acids) and form two distinct
RNA-binding surfaces, often called the proximal and
distal surfaces, respectively (9). These two RNA-binding
surfaces are used to promote sRNA–mRNA base pairing
by increasing the local concentration of the two binding
partners. Bacterial strains lacking Hfq exhibit increased
sensitivity to stress and reduced pathogenesis among
other phenotypes (10–14). One of the most intriguing
features of Hfq-mediated gene regulation is the network
of regulatory responses that allows a single sRNA to
trigger a multi-dimensional response to stress (15–17).
For instance, the sRNA RyhB regulates multiple genes
(including sodB, ftnA, bfR, acnA and sdhC) to help the
organism respond to low iron concentration (18). While
2 Nucleic Acids Research, 2012
binding both the proximal and distal surfaces of Hfq.
Structural characterization of glmS revealed two potential
(ARN)x motifs similar to those found in fhlA and rpoS.
Using a GFP fusion system we show that one of the two
upstream (ARN)x elements is essential for Hfq-mediated
regulation of glmS by both GlmZ and GlmY whereas the
other is dispensable. Furthermore, we probed the requirement of the variable C-terminal region of Hfq for regulation—a topic that has been very controversial. We show
that the Hfq C-terminus is not required for the classical
Hfq regulation involving GlmZ pairing with GlmS.
However, at least seven residues of the C-terminal extension (residues 65–72) were essential for GlmY activation
of GlmS expression mediated through YbhJ and GlmZ.
Thus the C-terminal extension may be involved in systems
where ancillary RNA or protein partners (such as YhbJ)
must be recruited to the complex during regulation.
MATERIALS AND METHODS
DNA oligonucleotides
The complete list of DNA oligonucleotides used for
cloning is provided as Supplementary Table S1.
Bacterial strains, media and growth conditions
The E. coli strain Top10 (Invitrogen) was used for cloning,
GFP fusion expression analysis and in all that involved
coexpression of GFP fusions with sRNAs and Hfq
variants. The hfq strains were also derived from Top10
cells and were generated as described previously by
replacing hfq with chloramphenicol or kanamycin resistance cassettes using the Quick and Easy Conditional
Knockout Kit (Gene Bridges) (33). In order to comply
with the usage of antibiotics, the chloramphenicol resistance cassette was removed from the hfq::CamR strain as
described in the Quick and Easy Conditional Knockout
Kit (Gene Bridges) operational manual. The successful
removal of the CamR cassette was verified by antibiotic
screening followed by PCR.
Growth in Luria–Bertani (LB) broth or on LB plates at
37 C was used throughout this study. Antibiotics were
applied at the following concentrations: 100 mg/ml ampicillin, 100 mg/ml spectinomycin, 34 mg/ml chloramphenicol and 30 mg/ml kanamycin.
Plasmids
Fusion plasmids
All GFP fusion systems were constructed starting from the
plasmid pBacEmGH [gift of the Cunningham Lab (34)]. In
brief, a glmS mRNA fragment from 162 to+21 relative to
the translation start site was cloned behind a pBAD
promoter to prepare pNS9006 (Supplementary Figure
S2). Primers NS20 and NS21 were used to amplify the
glmS fragment from genomic DNA and inserted into
pBacEmGH using NotI and NheI restrictions endonucleases. The plasmid was then verified using DNA sequencing.
Preparation of ARN mutants in glmS was carried out by
synthesizing glmS using primer extension where mutations
were introduced by the primers that were used. To
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the use of Hfq in many other stress pathways such as cold
shock (DsrA) (19), oxidative stress (OxyS) (20) and sugar
stress (SgrS) (21,22) are well established, a common mechanism describing how Hfq controls all these simultaneously is lacking.
The recent discovery of Hfq-binding elements, so-called
(ARN)x motifs, found in upstream regions of those
mRNAs that are regulated by sRNAs has provided new
clues for how Hfq helps mediate gene regulation. In both
rpoS and fhlA mRNAs, these Hfq-binding sequences are
essential for regulation (23–25) and structural data confirmed that the (ARN)x motif specifically interacts with
the distal RNA-binding surfaces of Hfq hexamers (26). If
these upstream Hfq-binding motifs are a common feature
of regulated messages, this could help explain how Hfq
rapidly targets specific mRNAs for regulation during
stress. In theory, as described in the fhlA work, Hfq could
pre-form complexes with relevant mRNAs and could
promote an efficient stress response upon sRNA synthesis
if and when it is induced by environmental stimuli (25).
Furthermore, recent work on the DsrA-rpoS system has
shown that Hfq binding to rpoS refolds the mRNA for a
favorable interaction with the sRNA (27). These observations, although currently limited to a few systems, help
explain the complex manner by which Hfq facilitates
integrated responses to environmental stress.
To further understand the importance of upstream
(ARN)x motifs in Hfq-dependent gene regulation, the
glmS system was studied. glmS encodes an essential
enzyme glucosamine-6-phosphate synthase (GlcN-6-P)
(28). In many Gram-positive bacteria (such as Bacillus
subtilis) the expression of GlmS is tightly regulated by a
self-cleaving ribozyme located within glmS that catalyzes
cleavage in response to high GlcN-6-P concentrations (29).
However, in Gram-negative species, GlmS expression is
regulated by two sRNAs GlmZ and GlmY, a dual regulatory system that requires Hfq for activity (30). In addition
to Hfq and RNase E, YhbJ, a predicted NTPase, was
shown to be involved in this regulatory network in
Escherichia coli (31) (Figure 1). In brief, GlmS is encoded
as a poly-cistronic message that also includes GlmU, a gene
for a bifunctional enzyme that produces UDP-GlcNAc. In
order to individually regulate GlmS expression, RNase E
cleaves the glmU–glmS transcript at the stop codon of
glmU to produce a monocistronic glmS transcript that is
translationally repressed by its 50 -UTR structure (30).
GlmZ interacts with glmS to unmask the ribosomebinding site (RBS) and promote translation. Additional
modulation is provided by YhbJ, a protein involved in
GlmZ turnover and thus negatively regulates GlmS expression by decreasing the amount of GlmZ available to
activate glmS (30). GlmY is a second sRNA involved in
this pathway. GlmY has significant sequence and structural homology to GlmZ such that when it is expressed,
GlmY acts as a decoy, recruiting YhbJ away from GlmZ
(32). Thus, GlmY prevents GlmZ degradation increasing
GlmZ concentration and allowing more glmS-GlmZ
adduct formation, with a net result of enhancing GlmS
expression.
In this work, we show that the glmS interaction with
Hfq is similar to what was previously reported with fhlA,
Nucleic Acids Research, 2012 3
construct pNS 9009 that contains the ARN 1 mutation in
glmS, five cycles of PCR were performed using primers
NS23, NS24 and NS27 where the desired mutation was
introduced in NS24. The PCR amplification was performed
using Taq polymerase according to standard procedure.
After the first five cycles, the PCR product was diluted
100-fold into a fresh PCR reaction to amplify the 50 - and
30 -regions of glmS using primers NS22 and NS21. The
amplified PCR fragment was then subjected to NotI/NheI
restriction digestion before cloning into pBacEmGH.
Similarly pNS9010 (derivative of pNS9006::ARN2,
prepared using NS23, NS25 and NS27) and pNS9011
(derivative of pNS9006::ARN12, prepared using
NS23, NS26 and NS27) were constructed. Successful
construction of the plasmid was validated by DNA
sequencing.
sRNA plasmids
sRNA plasmids, pNS9007 (GlmZ) and pNS9008 (GlmY)
were based on pNM12 (35). sRNA fragments for GlmZ
(NS37/NS38) and GlmY (NS35/NS36) were amplified
using DNA from TOP10 E. coli cells. The amplified fragments were then inserted into pNM12 after restriction digestion using MscI/EcoRI. The plasmids were verified by
DNA sequencing.
Hfq plasmids
All
Hfq
constructs
were
based
on
the
plasmid ptac-KanL94P (Addgene (34) plasmid 15927)
(36). Hfq wild-type (pNS9012, NS42/NS43), Hfq65
(pNS9013,NS42/NS44), Hfq72 (pNS9014, NS42/NS45),
Hfq87 (pNS9015, NS42/NS46) were amplified using
Top10 E. coli genomic DNA. Hfq mutants Y25A Hfq
(pNS9017), K56A Hfq (pNS9020) and H57A Hfq
(pNS9016) were amplified using a plasmid as template
that was prepared previously (9). Clostridium perfringens
Hfq (pNS9018, NS47/NS48) and Clostridium difficile Hfq
(pNS9019,
NS49/NS50)
were
amplified
using
C. perfringens and C. difficile genomic DNA. The PCR
fragments were then restriction digested with BamHI and
HindIII and inserted into ptac-KanL94P. The plasmids
were verified by DNA sequencing.
Fluorescence data collection
GFP expression of fusion constructs was measured after
lysis of cells grown to early stationary phase. All cultures
were grown at 37 C in LB media with appropriate antibiotics at concentrations listed above. Bacterial cultures
were generated by diluting overgrown overnight cultures.
After 3 h of growth, the cultures were induced with 0.1%
of arabinose. Cells were then allowed to grow for another
2.5 h. For Hfq complementation studies using the
pTac-plasmid system, Hfq expression was induced using
1 mM IPTG after 1 h of growth of diluted cultures before
inducing the expression of GlmS-GFP fusion proteins at
3 h. Cells were grown for an additional 2.5 h, before the
absorbance was measured at 600 nm to monitor the
growth of cells. 3.0 ml of cells were then harvested by
centrifuging at 10 000 rpm for 4 min. The cells were then
resuspended in 200 ml of lysis buffer (50 mM Tris–HCl pH
7.5, 25 mM NaCl, 2 mM EDTA). To lyse cells 15 ml of
lysozyme (20 mg/ml, Fisher), 30 ml of protease inhibitor
solution (complete EDTA-free protease inhibitor tablet
dissolved in 8 mL Roche) and 30 ml of 1% TritonX-100
was added followed by an incubation of 30 min at 37 C
while shaking. The lysed cells were then centrifuged
briefly. An amount of 200 ml of the supernatant was
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Figure 1. Mechanism of Hfq-mediated regulation of GlmS with sRNAs, GlmZ and GlmY [adapted from (31)]. glmS is transcribed as a
poly-cistronic message that also includes glmU. Upon transcription, RNase E cleaves at the stop codon of glmU (at 161 nt position with
respect to the translation start site of glmS) that separates the translational control of GlmS and GlmU genes. The RNase E cleaved glmS transcript
is repressed for translation by its 50 -UTR structure that masks the RBS. Upon synthesis of GlmZ sRNA, GlmS translation is activated in an
Hfq-dependent manner. Here the GlmZ–glmS interaction relieves the inhibitory structure of glmS for translation. YhbJ, a predicted NTPase
negatively regulates GlmS expression by processing GlmZ sRNA. GlmY, a second sRNA which has structural and sequence homology to GlmZ,
antagonizes YhbJ from GlmZ, upregulating GlmS expression. Overall, the GlmS expression is activated by two homologous sRNAs GlmZ and
GlmY in a coordinated manner in the presence of Hfq.
4 Nucleic Acids Research, 2012
then withdrawn to measure the fluorescence intensities of
GFP in a 96-well flat bottom black plate (CorningÕ ).
Fluorescence measurements were made at an excitation
wavelength of 485 nm and at an emission wavelength of
535 nm using a Tecan GENios Plus multi label plate
reader. The detector gain values were consistently set
between 95 and 103. All fluorescence measurements were
normalized to an identical culture where the expression of
GFP was not induced by arabinose to account for cellular
fluorescence. All measurements were at least in triplicate
and error values for each experiment were based on the
standard deviation between independent trials.
footprinting of Hfq on the folded RNA has been described
previously (25,37,39).
Western blot analysis
Total cellular extracts were collected from appropriate
strains and were lysed using sonication. Total protein
was separated on 15% (w/v) polyacrylamide gels
and electroblotted to a PVDF membrane. The
membrane was probed with rabbit anti-Hfq polyclonal
anti-sera. The antigen–antibody complex was visualized
using goat anti-rabbit immunoglobin horseradish peroxidase conjugated antibody (Super Signal West Pico
Chemiluminescent Substrate, PIERCE).
RNA preparation for SHAPE and gel shift analysis
Electrophoretic mobility shift assays
All binding reactions were performed in 50 mM Tris–HCl
pH 7.5,100 mM KCl and 10 mM MgCl2 at room temperature. Prior to any interactions all RNAs were annealed at
90 C for 3 min in buffer, and cooled to room temperature
for 30 min. For all reactions 10 ml aliquots were loaded after
diluting with loading buffer (10% (w/v) sucrose, xylene
cyanol, bromophenol blue) under a power of 5 W on
native 5–7% polyacrylamide (37:1) gels in 1 TBE. Dried
gels were visualized by phosphorimaging using a Typhoon
9210 imaging system (GE LifeSciences). Quantification was
done using ImageQuant 5.1 (Molecular Dynamics) and
Kaleidagraph 3.0 (Synergy). Data were fit using nonlinear
least-square analysis to a cooperative binding model shown
below [Equation (1)]. Here, L is the ligand concentration
and the cooperatively is indicated by n. Typical values for
n ranged from 1.5 to 2.1.
QFraction ¼
ðLÞn
KD+Ln
ð1Þ
In the case of A18, DsrA competition assays, the
glmSHfq complex was pre-formed and A18 and DsrA
was titrated at varying concentrations from 0 to 30 mM.
Chemical SHAPE analysis and footprinting
The method for using SHAPE technology to determine
the secondary structure of an mRNA 50 -UTR and the
RESULTS
glmS interaction with Hfq
Previous work performed on the glmS regulatory system
in E. coli showed the requirement of Hfq for GlmZ- and
GlmY-mediated activation of glmS (31,32). While the
basic outline of the regulatory network was established,
no molecular level understanding was developed. Based
on our previous work with OxyS/fhlA (25) and DsrA/
rpoS (24) we asked whether Hfq pre-binds the glmS
mRNA and how similar the glmS–Hfq interaction is
relative to other systems. The glmS mRNA fragment
used throughout this study was 182 nt in length. This
segment extends from position 161 nt upstream of the
GlmS start codon, which is the RNase E processing site
that separates the glmU–glmS bicistronic transcript
(Figure 1), and extends +21 nt into the coding region.
We used this construct to test whether glmS behaves in
a manner similar to fhlA (25), and to determine whether
one or both RNA-binding surfaces were involved in the
interaction with Hfq (Figure 2).
To measure the binding constant between glmS and
Hfq, [50 -32P] glmS mRNA was incubated with varying
Hfq concentrations from 0 nM to 2 mM (hexamer). The
glmS–Hfq complex was resolved on native gels to
provide a KD of 30 nM (Figure 2A). This value was
comparable to the interaction between fhlA–Hfq and is
typical of RNA–Hfq interactions measured previously
(9,24,25,40).
To determine which binding surfaces of Hfq interact
with Hfq, a competition gel shift assay was carried out
where the glmS*–Hfq complex was pre-formed and
competed against bona fide distal (A18 RNA) and
proximal (DsrA) site binders of Hfq. As observed in
Figure 2B, the glmS*–Hfq complex could only be
displaced by the simultaneous addition of both DsrA
and A18 showing that GlmS interacts with both faces of
Hfq similar to fhlA and rpoS (25).
Secondary structure of glmS mRNA and Hfq-binding
sites probed using SHAPE
SHAPE (37,41) was carried out to determine the secondary structure of glmS mRNA to identify its functional
elements. Figure 3 depicts the secondary structure for
glmS mRNA derived from SHAPE constraints,
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A18 RNA was purchased from Thermo Scientific
Dharmacon Technologies (Lafayette, CO) and deprotected following the manufacturer’s protocol. RNA
quality was assessed using denaturing PAGE and gel
purified. For in vitro binding analysis, glmS mRNA was
transcribed using a DNA fragment that amplifies the glmS
from 161 to +21 in E. coli XL-10 cells using primers
NS33 and NS34 and digested with Ssp I prior to transcription. For SHAPE analysis, glmS mRNA was in vitro
transcribed using a DNA template that was amplified
from XL-10 cells using primers NS31 and NS32, that
includes a structure cassette in the 50 - and 30 -regions as
previously described (37). In vitro transcription was performed after digesting the amplified product with Ssp I.
DsrA was obtained by runoff transcription of
pBAU10301 that was digested by Ssp I (38).
Nucleic Acids Research, 2012 5
superimposed with the modification intensity data. The
functional elements in glmS mRNA such as the potential
(ARN)x motifs, GlmZ-binding site, RBS and translation
start sites are also depicted in Figure 3. According to the
proposed glmS structure, two potential (ARN)x motifs
can be identified as candidates that could serve as
upstream Hfq-binding sites. To confirm Hfq-binding
regions in glmS, SHAPE footprinting was used as
described previously (25) and the results are indicated in
Figures 3 and Supplementary Figure S1. As expected,
both (ARN)x regions footprinted with Hfq among other
probable bindings sites indicated in Figure 3.
Measuring translational regulation of GlmS using a GFP
fusion system
A GFP fusion system was used to measure the
Hfq-dependent translational regulation of GlmS by
sRNAs GlmZ and GlmY. A schematic diagram of the
fusion expression system is shown in Supplementary
Figure S2. As described above, a glmS fragment
spanning the region of 161 to +21 was N-terminally
fused to GFP under the transcriptional control of a
pBAD promoter. The fusion system is constructed in a
pBAC vector backbone (kindly provided by the Philip
Cunningham Lab, Wayne State University, Biological
Sciences) which is based on the F-plasmid ori2/RepE
replicon, that allows stable maintenance of large
genomic fragments as very low or single copy plasmids
(42). In addition, the vector also contains an oriV replication origin that can be induced by L-arabinise to produce
high copy numbers of the plasmid in an oriV specific
manner when expressed in a host strain that can supply
the replication initiation protein TrfA in trans (43).
However, throughout this study a low copy number of
fusion plasmids were maintained since TrfA is not
present in these strains.
To monitor expression of GlmS in the presence of
sRNAs, GlmZ or GlmY were coexpressed together with
the GlmS:GFP reporter. Here vectors pNS9007 and
pNS9008, derivatives of the pBAD plasmid series, were
used to express GlmZ and GlmY. Therefore coexpression
of the GFP fusion and sRNAs were both under the
control of a pBAD promoter induced with 0.1% arabinose. The coordinated transcription of mRNAs and their
cognate sRNAs is required for efficient Hfq-mediated
regulation in vivo (15,16).
To confirm the proper functioning of the fusion construct and coexpression conditions, GlmS:GFP expression
was measured to obtain previously validated observations
on the GlmS regulatory system (31,32) and the results are
shown in Figure 4. To measure GFP fluorescence, liquid
cultures were grown from overgrown overnight cultures.
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Figure 2. glmS interaction with Hfq. (A) Analysis of glmS interaction with Hfq (left). Gel shift experiment showing the binary complex between Hfq
and glmS mRNA. 32P-labeled glmS* was titrated with varying concentrations of Hfq ranging from 0 to 2 mM. Quantitative analysis of the glmS–Hfq
gel (right). As described in materials and methods, thermodynamic constants were determined by non-linear least squares analysis fitted to a
cooperative binding model. (B) Competition gel-binding experiment to determine the Hfq-binding surface to glmS. The glmS*–Hfq complex was
pre-formed and incubated with increasing amounts of either distal (A18 RNA), proximal (DsrA) or both (A18 and DsrA) binding RNAs (0–30 mM) to
compete glmS* away from Hfq.
6 Nucleic Acids Research, 2012
New cultures were then grown at 37 C for 3 h while
shaking before inducing with 0.1% L-arabinose. The
cultures were left to grow for another 150 min upon induction before the cells were harvested. The growth of
each individual culture was monitored by its OD600
absorbance. Cells were consistently collected at their
early stationary phase throughout (OD600 1.4 ± 0.2).
Lower dilutions from overnight cultures were made for
slow growing strains, such as Dhfq cells to reach early
stationary phase within the time frame of the assay.
Fluorescence was measured upon lysis of 3.0 ml of cell
culture (see ‘Materials and Methods’ section). The fluorescence data measured from liquid cultures are shown in
Figure 4, where GFP expression is normalized to an identical un-induced culture to correct for cellular fluorescence
and used to calculate the fold expression of GlmS.
Basal levels of GlmS expression in the absence of either
sRNA are 2-fold that of an identical culture that was not
induced with arabinose. The presence of either GlmZ or
GlmY leads to an additional 2- to 3-fold increase in
GlmS:GFP expression. The ability to upregulate GlmS
expression is lost in the absence of Hfq, even in the
presence of sRNAs. These data correlate nicely with
previous observations of this regulatory system (31,32).
They show that the GFP fusion construct and the assay
conditions are able to capture the effects of sRNAs and
Hfq-mediated regulation of GlmS.
To test whether the translational fusion can recognize
defective Hfq species that are incapable of regulating gene
expression, proximal and distal mutants of Hfq were used
to measure GlmS activation (Figure 5). Y25A Hfq and
K56A Hfq mutants have been shown previously to
diminish nucleic acid binding at distal and proximal
RNA-binding sites, respectively, (9,25) and the H57
proximal residue has been implicated to be crucial for
hexamer organization in Pseudomonas aeruginosa (44)
and Salmonella typhimurium (45). Data presented in
Figure 5, indicates that either proximal (K56A) or distal
(Y25A) mutants are incapable of facilitating GlmS
upregulation in the presence of sRNAs. However, H57A
Figure 4. Expression of GlmS:GFP measured using GFP fluorescence.
Relative expression levels of GlmS are measured in response to sRNAs
GlmZ, GlmY and Hfq. Fluorescence was measured from E. coli cells
that were grown in liquid media according to the conditions described
in materials and methods. The growth of cells was controlled and
monitored using the cell density (OD600) where all cultures were harvested upon reaching early stationary phase (OD600 1.4 ± 0.2). Cells
were lysed before fluorescence was measured using a Tecan GENios
Plus multi label plate reader. All fluorescence measurements were at
least triplicated using independent inoculations. To calculate relative
expression levels of GlmS, the fluorescence levels of each sample was
compared to an identical uninduced culture to correct for cellular fluorescence. GlmS:GFP and sRNA synthesis was controlled using a pBAD
promoter inducible by L-arabinose. To measure the Hfq dependence on
GlmS expression, experiments were carried out in a Dhfq background.
Hfq shows a weak facilitation towards GlmS expression
compared to wild-type Hfq. These data indicate the
proper functioning of the GlmS:GFP fusion system, and
that it is sensitive to sRNAs (GlmZ/GlmY) and Hfq.
Upstream Hfq-binding (ARN)x elements are essential
for regulation
(ARN)x motifs in upstream regions are essential for Hfqmediated gene regulation of fhlA (25) and rpoS (23,24).
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Figure 3. SHAPE-derived secondary structure model for glmS mRNA. The NMIA reactivity for each nucleotide corresponds to a given color,
nucleotides with reactivities >0.6 are shown in red, from 0.3 to 0.59 in purple and <0.3 in grey. Hfq footprinting data in the presence and absence of
0.5 mM Hfq hexamer are superimposed on the structure. Large wedges indicate where there was a strong protection (>0.6) from wild-type Hfq
observed and small wedges where a weak protection (0.3–0.59) was observed. The (ARN)x sites, glmZ-binding site, RBS and the start codon are also
shown.
Nucleic Acids Research, 2012 7
Requirement of the C-terminal domain of Hfq for
regulation of GlmS by sRNAs GlmZ and GlmY
Structural evidence also indicates the preferential binding
of (ARN)x motifs at the distal site of Hfq (26).
Furthermore, a recent study showed the specific enrichment of these sequence elements in genomic SELEX
experiments that were performed to identify Hfq binders
(46). The above observations lead to a model where, the
presence of upstream (ARN)x motifs could be a common
occurrence in Hfq-dependent mRNAs. The secondary
structure model together with Hfq footprinting data for
glmS, indicated the presence of two potential (ARN)x
motifs either of which could be required for regulation
(Figure 3). The GFP fusion system was used to identify
(ARN)x element(s) that are essential for GlmS activation
in vivo. Both potential ARN motifs were mutated serially
(glmS 1 and glmS 2) and simultaneously (glmS 12)
to identify the role of each in glmS regulation (also see
Figure 3).
The expression of GlmS 1:GFP, GlmS 2:GFP and
GlmS 12:GFP was monitored in the presence of
GlmZ (Figure 6A) and GlmY (Figure 6B). Data in
Figure 6A show that, sRNA GlmZ is unable to
activate GlmS expression of the glmS 2 mutant.
However GlmS expression is higher than wild-type
levels for the glmS 1 mutant. The overactivation
caused by glmS 1 was absent when both ARN motifs
were mutated (12). To test whether glmS 1 activation is Hfq dependent, the regulation was monitored in
the absence of Hfq. Data from this experiment reveals
that the 1 mutation did not change the Hfq-dependence
of GlmS activation by GlmZ. However, neither ARN
mutant (1 nor 2) were able to upregulate GlmS expression in the presence of GlmY (Figure 6B), implying
that the 1 mutation only affects the GlmZ pathway.
These results indicate the requirement of (ARN)x
elements for Hfq-mediated regulation of GlmS and that
ARN-2 is the most essential Hfq-binding motif for both
GlmZ and GlmY pathways.
Downloaded from http://nar.oxfordjournals.org/ at Wayne State University on June 4, 2012
Figure 5. GlmS:GFP expression levels in the presence of mutant Hfq
variants. To further characterize the functionality of the fusion system,
GlmS expression levels were monitored in the presence of sRNAs
with mutant Hfq variants that were previously shown to deter Hfq
function. External Hfq for these experiments was supplemented using
a plasmid system where the synthesis of Hfq was under the control of a
pTac promoter that is inducible by IPTG. Here Hfq expression was
induced prior to GlmS:GFP and sRNA synthesis to allow efficient
regulation.
Hfq is present in most Gram-positive and Gram-negative
bacteria. In all cases the nucleic acid-binding Sm1 and
Sm2 motifs are evolutionary conserved across species
which spans positions 7 through 66 based on the E. coli
numbering. The greatest variability is observed at the
C-terminus of Hfq where E. coli and Salmonella enterica
have fairly long tails (102 amino acids) while species
such as C. difficile and C. perfringens have shorter
C-terminal extensions. The functional importance of the
C-terminus of Hfq for regulation is unclear and conflicting
reports have been presented previously (47–49). Therefore,
the necessity of the C-terminal region of Hfq for GlmS
activation by sRNAs, GlmZ or GlmY was tested using the
GFP fusion system.
Several C-terminal truncations of E. coli Hfq were
coexpressed together with the GlmS:GFP fusion protein
and sRNAs in a Dhfq background to identify if the
C-terminal amino acids are required for regulation. In
addition to wild-type Hfq that consists of 102 amino
acids, shortened C-terminal constructs of Hfq87 (87
amino acids), Hfq72 and Hfq65 were prepared. Here the
Hfq65 variant can essentially be regarded as the tail-less
version of Hfq since it only consists of the core Sm1 and
Sm2 motifs. All C-terminal variants of Hfq were constructed using the ptac-KanL94P plasmid as the parent
vector (36), that is under the transcriptional control of
an IPTG inducible pTac promoter. The assay conditions
to measure the GFP expression were similar to those
described earlier, although Hfq induction was carried
out using 1 mM IPTG, 1 h before arabinose induction to
allow synthesis of Hfq prior to sRNA regulation. Relative
expression levels of GlmS were monitored as the ability to
upregulate GlmS synthesis individually by sRNAs GlmZ
and GlmY with varying C-terminal extensions of Hfq
(Figure 7).
From data shown in Figure 7, it is clear that plasmid
borne and native Hfq (Hfq wt) regulate GlmS expression
at comparable levels regardless of whether GlmZ or GlmY
is used as the activator. GlmS regulation by GlmZ was
mostly independent of the length of the Hfq C-terminal
extension as Hfq72 and Hfq87 showed identical levels of
GlmS expression to wild-type Hfq. Hfq65 on the other
hand showed reduced levels of GlmS expression
implying a possible effect caused by shortening the Hfq
C-terminal tail. Semi-native western blot analysis of
cellular Hfq65 showed multiple Hfq conformations
relative to wild-type Hfq implying possible destabilization
of the Hfq functional core in the absence of C-terminal
residues (Supplementary Figure S3). However, all Hfq
C-terminal variants (including Hfq65) were better
at upregulating GlmS expression than a Dhfq strain in
the presence of GlmZ, indicating at least some functional
hfq hexamer is present in vivo (Figure 7). Conversely, for
GlmY-mediated activation, Hfq65 was unable to
upregulate GlmS above basal levels while Hfq72 and
Hfq87 were proficient. These data show that the absence
of amino acid residues from at least 66–72 affected
GlmY’s ability to upregulate GlmS in a manner distinct
8 Nucleic Acids Research, 2012
Figure 7. Requirement of Hfq C-terminal domain for GlmS:GFP
upregulation by GlmZ and GlmY. Activation of GlmS:GFP expression
by sRNAs was measured in the presence of Hfq that consists of varying
C-terminal extension lengths. In addition to Hfq wild-type (102 amino
acids), Hfq65 (65 amino acids), Hfq72 and Hfq87 were tested for its
ability to perform regulation. Hfq65 consists only the nucleic
acid-binding Sm1 and Sm2 domains while Hfq72 and Hfq87 have 7
aa and 22 aa extensions off the Sm cores. All externally supplemented
hfq was under the transcriptional control of a pTac promoter that was
induced using 1 mM IPTG. Coexpression and induction conditions of
GlmS:GFP, sRNA and Hfq plasmid systems are described in material
and methods.
from GlmZ regulation. However, as described above,
GlmZ and GlmY act in a hierarchical manner during
regulation. Therefore a loss of GlmY’s ability to recruit
YhbJ for self-processing would affect GlmZ’s ability to
activate GlmS synthesis. These data provide sufficient
evidence to suggest the importance of at least the first
seven amino acids of the C-terminal tail of Hfq for some
in vivo functions.
Clostridium perfringens and C. difficile Hfq variants are
proficient in GlmZ but not GlmY-mediated regulation
of GlmS in E. coli
To validate these observations that the first 7-amino acids
of the C-terminal tail of Hfq were essential for the GlmY
pathway, Hfq variants belonging to Gram-positive bacteria
C. difficile and C. perfringens were used to substitute for
E. coli Hfq to activate GlmS synthesis. Both HfqCd and
HfqCp variants have similar nucleic acid-binding regions
(Sm1/Sm2) compared to E. coli. However, not much
sequence homology is observed beyond the Sm regions
(Figure 8A). Therefore, the nucleic acid-binding properties
should be conserved in HfqCd and HfqCp to promote
GlmZ–glmS base pairing. However, if there is a need for
the C-terminal extension to facilitate regulation, HfqCp and
HfqCd may not be able to cross complement GlmY native
functions.
GlmS activation was monitored in the presence of HfqCd
and HfqCp variants with either GlmZ or GlmY sRNAs in a
Dhfq background. The results are shown in Figure 8B.
HfqCd and HfqCp both successfully promote GlmZdependent GlmS activation. In fact, increased upregulation
of GlmS is observed for HfqCp in the presence of GlmZ.
These data fundamentally confirm that the GlmZ–glmS
interaction, which revolves around the ability of Hfq to
facilitate base-paring between a sRNA and its target
mRNA, can be achieved so long as the core nucleic acidbinding domain is present. However, HfqCd and HfqCp
were unable to upregulate GlmS expression via the GlmY
pathway. Gel shift assays confirmed that GlmY binds
HfqEc and HfqCp, although its affinity for HfqCp is about
10-fold weaker than for HfqEc. (Supplementary Figure S4).
In contrast, GlmZ binds HfqCp and HfqEc with only a slight
loss of affinity implying some differences in the manner
with which GlmZ and GlmY bind Hfq despite their
sequence and structural homology. Since the only significant difference between the three Hfq variants are at
the C-terminal region, a potential role of the C-terminus
is implied for GlmY activation, where additional
protein–protein or protein–RNA interactions are likely
required.
DISCUSSION
Being able to adapt to environmental stress is vital for the
survival and propagation of bacteria. The RNA-binding
Downloaded from http://nar.oxfordjournals.org/ at Wayne State University on June 4, 2012
Figure 6. Effect of upstream Hfq binding (ARN)x elements in GlmS activation by GlmZ and GlmY. (A) Relative GlmS:GFP expression levels of
glmS ARN mutants measured in the presence of GlmZ. Here mutations to glmS ARN elements were made as indicated in above Figure 2. Since two
potential ARN motifs were detected they were removed sequentially and simultaneously to produce glmS 1, glmS 2 and glmS 12 respectively.
The expression level of GlmS was measured as indicated in material and methods. The relative upregulation of GlmS was compared to expression
levels when in the absence of GlmZ and Hfq. (B) Upregulation of glmS ARN mutants were measured with GlmY and compared to wild-type glmS
expression levels.
Nucleic Acids Research, 2012 9
protein Hfq plays a key role in modulating stress responses in bacteria by regulating gene expression in a
sRNA-dependent manner. Hfq functions primarily in
promoting sRNA–mRNA base pairing by increasing the
local concentrations of sRNAs and target mRNAs
through its distal and proximal RNA-binding surfaces.
In addition, Hfq acts as a chaperone to trigger structural
transitions in RNAs to initiate recognition between
cognate sRNA–mRNA pairs (27), or partners directly or
indirectly with processing enzymes such as RNase E
(50,51), Poly A polymerase (52), PNPase (50), RNA
helicases (53,54), RelA (55) and YhbJ (30) to aid stringent
gene regulation. These diverse functions enable Hfq to be
involved in a cascade of regulatory events that control
multiple stress responsive pathways. Since all Hfq
networks involve sRNAs to initiate gene regulation at
the post-transcriptional level, the sRNA–mRNA interaction can be regarded as the main trigger for downstream
regulatory outcomes.
A number of recent reports have indicated that
upstream (ARN)x sequence motifs are common among
Hfq-dependent mRNAs and help recruit Hfq to these
mRNAs (23,25,56). Structural evidence shows that
(ARN)x motifs interact with the distal RNA-binding site
of Hfq while the proximal site harbors sRNAs during
regulation (26). In this work, (ARN)x motifs were
identified and characterized within glmS mRNA, a
message regulated by sRNAs GlmZ and GlmY in an
Hfq-dependent manner. The interaction between Hfq
and glmS is similar to what was previously observed for
fhlA (25) where the mRNA uses both RNA-binding
surfaces to interact with Hfq in the absence of sRNAs.
This mode of interaction between mRNAs and Hfq was
attributed to a model wherein, the complexes could serve
as a precursor RNP complex that enables a rapid response
to stress in bacteria.
Examining the proposed secondary structure of glmS
and Hfq footprinting data obtained using SHAPE, it
was apparent that two potential ARN motifs (ARN-1
and ARN-2) were present upstream of the RBS
(Figure 3). To identify which ARN element(s) were essential for regulation, glmS mRNA was fused N-terminally
to GFP to measure GlmS expression levels. GlmS translation was upregulated in the presence of either sRNA in
an Hfq-dependent manner confirming the proper functioning of the fusion system. Of the two potential ARN
motifs, ARN-2 had the most adverse effects on regulation in the presence of either sRNA. However, the loss of
ARN-1 caused a hyper-activation of GlmS only in the
presence of GlmZ. Nonetheless, upregulation of glmS
ARN-1 was Hfq dependent, implying that the regulatory node was still intact despite the changes in glmS functional elements. These results indicate that ARN-2 is the
essential (ARN)x motif in glmS required for Hfq-mediated
regulation. While we do not completely understand why
glmS-ARN-1 led to over-activation, it might result from
structural changes in glmS due to the sequence modifications required to ablate the Hfq binding site.
Alternatively, it could also be explained by a partitioning
model. If Hfq must be present at ARN-2 for glmZ activation, and Hfq can bind at either ARN-1 or ARN-2, destruction of ARN-1 would lead to an increased population
of the active adduct (Hfq-binding to ARN-2) triggering a
more favorable response than wild-type glmS. The key
Downloaded from http://nar.oxfordjournals.org/ at Wayne State University on June 4, 2012
Figure 8. GlmS:GFP upregulation using C. perfringens and C. difficile Hfq variants by GlmZ and GlmY. (A) Sequence homology between E. coli,
C. difficile and C. perfringens Hfq variants. Nucleic acid-binding Sm1/Sm2 domains and important amino acid residues in proximal and distal
RNA-binding surfaces required for regulation are indicated. (B) Cross-complementation for HfqEc using HfqCp and HfqCd homologues for GlmS
upregulation. The ability to activate GlmS expression by GlmZ and GlmY was measured when HfqCd and HfqCp was supplemented in E.coli Dhfq
cells.
10 Nucleic Acids Research, 2012
analysis of Hfq65 indicates some issues associated with
speciation or reduced stability of the functional Hfq
hexamer (Supplementary Figure S3) as previously
postulated (13). This result explains the reduced activity
of GlmZ-mediated upregulation of glmS expression in the
presence of Hfq65. However, the inability of GlmY to
utilize Hfq65 to any appreciable extent remains puzzling
since a significant amount of functional hexamer is still
present in these cells based on western blot analysis.
HfqCp and HfqCd were used to cross complement for
HfqEc in order to completely rule out the potential differences in Hfq utilizations by sRNAs during regulation. Hfq
variants from C. perfringens and C. difficile fully complemented the hfq strain when measured based on GlmZ
activation of GlmS. GlmY, however, was unable to take
advantage of cross-species complementation. Sm cores in
HfqCp and HfqCd are highly conserved compared to HfqEc
and successfully promote GlmZ–glmS base pairing that
represents the classical Hfq mediation process through
its nucleic acid-binding domains. The inability of GlmY
to use Hfq65, HfqCp and HfqCd to upregulate GlmS
suggests that the Sm core alone is insufficient for regulation in this pathway. Since the primary functional role of
GlmY is to self-direct the processing complex that utilizes
YhbJ and polyA polymerase (PAP I) (31), Hfq C-terminal
region may facilitate the assembly of this complex through
additional RNA and/or protein interactions with Hfq.
The Hfq C-terminal region has long been predicted as a
landing pad for either protein or nucleic acid-binding
partners. In fact, recent structural data compares the
Hfq C-terminal regions to intrinsically disordered
proteins (IDPs) that could potentially facilitate intermolecular interactions (47,57). Pull down experiments carried
out to map Hfq-protein complexes indicate that Hfq interacts directly or indirectly with proteins in the bacterial
RNA degradosome such as RNase E, polynucleotide
phosphorylase (PNPase), poly A polymerase (PAP I)
(59,60) and a variety of regulatory proteins in bacteria
(T. Lee and A. Feig, unpublished data). Furthermore, a
recent study by Prévost et al. proposed a model for the
physiological relevance of an RNase E–Hfq interaction
during RNA degradation triggered by the sRNA–
mRNA recognition (61,62). Since no detectable protein
interactions have been reported within the Sm cores,
these complexes should be accommodated at the
C-terminal region of Hfq. Whether Hfq recruits YhbJ as
part of a novel processing complex that affects sRNA
turnover or whether it promotes an RNA interaction
with GlmY through its C-terminal that is essential for
GlmY turnover needs to be tested further.
Overall, this work further illustrates the requirement for
Hfq to bind mRNAs through upstream (ARN)x motifs
during sRNA-dependent gene regulation. In addition,
these studies show that a non-canonical sRNA pathway
requires additional Hfq interactions traced to the
C-terminal extension that are not essential for the canonical sRNA–mRNA pairing behavior typically carried out
by Hfq. C-terminal amino acids 66–72 from E. coli are
essential for GlmY-mediated regulation of GlmS,
implying a conditional requirement for the C-terminal
region depending on the mode of sRNA recruitment and
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point however, is that, as observed in previous cases such
as in rpoS, rprA (23) and fhlA (25), Hfq-mediated regulation of glmS is absolutely dependent on the presence of an
upstream (ARN)x element, adding to the list of networks
that rely on such sequence motifs for function and may
indicate that this is a global requirement within mRNAs
that are destined for Hfq-dependent regulation.
The manner by which Hfq recognizes target mRNAs
could be important for understanding the mechanism of
stress regulation in bacteria. This interaction could allow
HfqmRNA complexes to form prior to stress, thus identifying those messages that might be regulated during a
time-sensitive response. While we have no definitive data
to confirm this hypothesis, this model would explain Hfq’s
ability to handle multiple regulatory networks using
multiple sRNAs. Recent evidence also suggests that Hfq
can be a limiting factor in cells and that a coordinate expression of sRNA and mRNAs is required for the proper
functioning of regulatory nodes (15). This implies the
intense competition between RNAs to acquire Hfq
which in turn explains the need for expressed mRNAs to
capture Hfq to facilitate their own regulation. Other functions of this interaction could include initiation of structural rearrangements in the mRNA to facilitate the
interaction with sRNA as was observed in the case of
DsrA and rpoS (27). However, the structural transitions
that are triggered by the mRNA–Hfq interaction alone are
insufficient to affect basal translation levels. The glmS–
Hfq interaction alone had no effect in GlmS expression,
for instance (data not shown). Similar observations were
reported for sodB, rpoS, ompC and fhlA, all having only
minimal effects on translation upon binding to Hfq (15).
The role of Hfq in stress regulation throughout the bacterial kingdom is well established. Comparison of Hfq
variants across species has confirmed the presence of an
evolutionary conserved nucleic acid-binding core (Sm1
and Sm2) and a variable C-terminal region that is less
well preserved. While the requirement of the Sm core for
Hfq function is clear, involvement of the C-terminal
region of Hfq in regulation is still controversial (49,57).
To gain more information regarding the function of the
C-terminal region of Hfq during regulation, the glmS
system was used. A clear advantage of using this system
to address this problem stems from the presence of two
regulatory layers that are directed by GlmZ and GlmY in
an Hfq-dependent manner. While the role of Hfq in
facilitating the GlmZ–glmS interaction is well established,
the requirement of Hfq by GlmY in YhbJ processing is
unclear. Since an interaction between GlmY and glmS is
not established, (31) YhbJ could be recruited by a
GlmY–Hfq complex potentially through the C-terminal
region of Hfq.
When the requirement of the C-terminal region of Hfq
for GlmS activation was measured, data indicated that
both Hfq72 and Hfq87 were equally proficient at assisting
GlmY and GlmZ upregulate GlmS. However, when only
the Sm core of Hfq was present (Hfq65), only GlmZ
allowed significant upregulation of GlmS expression
relative to a Dhfq strain. Although crystallographic data
show that C-terminal residues are not essential for
hexamer association (58), semi-native western blot
Nucleic Acids Research, 2012 11
may relate to additional RNA or protein-binding
interactions.
SUPPLEMENTARY DATA
Supplementary Data are available at NAR Online:
Supplementary Tables 1–2 and Supplementary Figures
1–4.
FUNDING
National Institutes of Health [GM075068 to A.L.F.,
in part]; Wayne State University, Office of the VicePresident for Research. Funding for open access charge:
NIH [GM-075068].
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